Skip to main content
NCBI home page
Applied and Environmental Microbiology logoLink to Applied and Environmental Microbiology
. 2016 Nov 21;82(24):7093–7101. doi: 10.1128/AEM.02572-16

Interactions of Plutonium with Pseudomonas sp. Strain EPS-1W and Its Extracellular Polymeric Substances

Mark A Boggs a,*,✉, Yongqin Jiao b, Zurong Dai a, Mavrik Zavarin a, Annie B Kersting a
Editor: F E Löfflerc
PMCID: PMC5118920  PMID: 27694230

ABSTRACT

Safe and effective nuclear waste disposal, as well as accidental radionuclide releases, necessitates our understanding of the fate of radionuclides in the environment, including their interaction with microorganisms. We examined the sorption of Pu(IV) and Pu(V) to Pseudomonas sp. strain EPS-1W, an aerobic bacterium isolated from plutonium (Pu)-contaminated groundwater collected in the United States at the Nevada National Security Site (NNSS) in Nevada. We compared Pu sorption to cells with and without bound extracellular polymeric substances (EPS). Wild-type cells with intact EPS sorbed Pu(V) more effectively than cells with EPS removed. In contrast, cells with and without EPS showed the same sorption affinity for Pu(IV). In vitro experiments with extracted EPS revealed rapid reduction of Pu(V) to Pu(IV). Transmission electron microscopy indicated that 2- to 3-nm nanocrystalline Pu(IV)O2 formed on cells equilibrated with high concentrations of Pu(IV) but not Pu(V). Thus, EPS, while facilitating Pu(V) reduction, inhibit the formation of nanocrystalline Pu(IV) precipitates.

IMPORTANCE Our results indicate that EPS are an effective reductant for Pu(V) and sorbent for Pu(IV) and may impact Pu redox cycling and mobility in the environment. Additionally, the resulting Pu morphology associated with EPS will depend on the concentration and initial Pu oxidation state. While our results are not directly applicable to the Pu transport situation at the NNSS, the results suggest that, in general, stationary microorganisms and biofilms will tend to limit the migration of Pu and provide an important Pu retardation mechanism in the environment. In a broader sense, our results, along with a growing body of literature, highlight the important role of microorganisms as producers of redox-active organic ligands and therefore as modulators of radionuclide redox transformations and complexation in the subsurface.

INTRODUCTION

The civil production of nuclear energy and military production of nuclear materials have resulted in an estimated worldwide inventory of over 2,000 metric tons of plutonium (Pu) (1). This inventory continues to increase at a rate of approximately 70 metric tons per year as a result of global nuclear energy production (2). Due to its long half-life (24,100 years for 239Pu) and high radiotoxicity (3), Pu is an important driver in public health risk assessments for nuclear waste repositories and radiologically contaminated sites. However, predicting how Pu behaves in the environment and ultimately calculating its human health risk are limited by our understanding of the dominant biogeochemical processes controlling its behavior.

The behavior of Pu in the environment is strongly dependent on its oxidation state and concentration. Among all the actinides, Pu has one of the most complex chemical, redox, and surface sorption behaviors. At low concentrations, Pu can exist as aqueous species in the III, IV, V, and/or VI oxidation states, all of which have different solubilities (4), mineral sorption affinities (5–7), and hence, environmental mobility and bioavailability. Under mildly oxic environmental conditions, the predominant oxidation states of aqueous Pu are the sparingly soluble Pu(IV) and the more soluble Pu(V) (8–10). The affinity of Pu(IV) to mineral surfaces is substantially greater than that of Pu(V), and this has been attributed to their difference in effective charge (11). However, it has been shown that Pu(V) can be reduced to Pu(IV) and even Pu(III) on a range of mineral surfaces, leading to a congruence in the apparent sorption behavior of Pu(IV) and Pu(V) (5). At high concentrations (>10−8 ± 1 M), Pu(IV) hydrolyzes to form Pu(IV) oxide precipitates (4, 12). This can lead to the formation of intrinsic Pu(IV)O2 colloids and/or surface precipitates (13).

Microorganisms are associated with a variety of radioactive materials at U.S. Department of Energy contaminated sites (14, 15), and it is apparent that Pu speciation and solubility can be affected by microbial activity (16–21). Previous studies of Pseudomonas stutzeri and Bacillus sphaericus reported a sorption affinity of nearly 100 μg Pu/gdry cell weight for Pu(VI) to the surface of vegetative cells and spores (20). Further investigations of Pu on B. sphaericus by extended X-ray absorption fine-structure (EXAFS) spectroscopy revealed that an insoluble Pu(VI) phosphate complex can form on the cell surface (22). Bacillus subtilis was shown to reduce surface-bound Pu(VI) to Pu(IV), with simultaneous reduction of Pu(VI) to Pu(V) in solution, indicating that cell-bound and soluble cell exudates are redox active with respect to Pu (23). Under anoxic conditions and in the presence of nitrilotriacetic acid (NTA), it was found that Bacillus sp. can also solubilize solid Pu(IV)O2(s) by reducing it to Pu(III) (24). Studies with the dissimilatory metal-reducing bacteria (DMRB) Geobacter metallireducens and Shewanella oneidensis reported the formation of inorganic Pu(IV) colloids after exposure to a mixture of Pu(V) and Pu(VI) under anoxic conditions (25) and the formation of Pu(III) in some cases (26–28). These laboratory studies demonstrate that Pu sorption to bacterial surfaces can be driven by a variety of mechanisms, including redox transformations, resulting in Pu products of different composition and stability. However, the cellular and/or extracellular components controlling these redox transformations and/or precipitation reactions under oxic conditions have not been examined in detail.

Among the known mechanisms that control metal transformation by microorganisms, complexation by extracellular polymeric substances (EPS) produced and secreted by microorganisms represents an important process. Previous research demonstrates that EPS of bacterial origin are able to interact with metals, including actinides such as uranium and neptunium, and alter their redox and solubility properties (29–32). However, to the best of our knowledge, no studies on the specific interaction of Pu with EPS have been reported under oxic conditions. Pu interaction with EPS is likely to induce redox transformations and, as a result, changes in Pu speciation and mobility. Compared to the reduction of Pu(IV) to Pu(III) that was observed under anoxic conditions (26, 28), facile Pu(V, VI) reduction to Pu(IV) is expected to occur under oxic conditions (25).

In the present study, we examined the sorption of Pu(V) and Pu(IV) to Pseudomonas sp. strain EPS-1W, an environmentally relevant aerobic bacterium collected, identified, and cultured from Pu-contaminated groundwater at the Nevada National Security Site (NNSS), formerly known as the Nevada Test Site, Nevada. To better understand Pu-cell interaction, we focused on the specific role of cell-bound EPS produced by the Pseudomonas sp. in controlling Pu sorption and redox transformation. Cell-bound EPS serve as the outermost layer of the cell surface and, as a result, are likely to play a role in Pu-cell interaction. We show for the first time that the cell-bound EPS of Pseudomonas sp., not the cells, control Pu(V) reduction and subsequent Pu(IV) sorption in synthetic near-neutral-pH, low-bicarbonate groundwater, similar to the composition of the groundwater at the NNSS. Our results provide insight into the role of microbes and bioproducts affecting the environmental mobility and fate of Pu.

MATERIALS AND METHODS

Pu stock preparation.

Unless otherwise noted, all chemicals were of high purity (ACS reagents) and used without any additional purification. 238Plutonium (98.9% 238Pu, 0.11% 241Pu, and 0.1% 239Pu by activity) and 242Pu (15.8% 238Pu, 5.1% 239 + 240Pu, and 79.1% 242Pu by activity) were used as Pu stock solutions. Prior to use, impurities were removed from the Pu stock solutions (in 8 M HNO3) by loading them onto a Bio-Rad AG 1-X8 resin and eluting the Pu as Pu(III) using an HI-HCl mixture. Excess HI was removed from the eluent by heating to dryness several times.

The Pu(IV) stocks were prepared by dissolving the purified 238Pu and 242Pu in 0.1 M HClO4 to yield 3 × 10−6 and 5 × 10−3 M Pu, respectively. The oxidation state was verified by UV-visible (UV-Vis) spectrometry and LaF3 precipitation (33) to be IV (>90%) (see the supplemental material for details). While LaF3 precipitation is a robust method of differentiating between the oxidized Pu(V, VI) and reduced Pu(III, IV) states of Pu, errors of 5 to 10% are inherent.

A 238Pu(V) stock was prepared by dissolving the purified 238Pu in 0.1 M HNO3 and adjusting to pH 3 with NaOH. Hydrogen peroxide, 0.05 M, was added and the stock heated gently (50°C), producing Pu(V). To remove any Pu(VI) remaining in solution, the stock was diluted in a buffer solution containing 0.7 mM NaHCO3 and 5 mM NaClO4 to achieve a circumneutral pH and then exposed to high-surface-area SiO2 (10 g/liter) to preferentially adsorb Pu(VI) (34). An LaF3 oxidation state analysis of the final 238Pu(V) stock (2 × 10−6 M) indicated that Pu(III, IV) constituted less than 6% of the total concentration.

A 242Pu(V) stock was prepared by applying a −1.95 V versus Ag/AgCl redox potential (Pine Instruments) for 1 week to the purified 242Pu (35), which oxidized Pu(IV) to Pu(VI). The 242Pu(VI) solution (verified by UV-Vis spectrometry) was then diluted 1:10 in 0.1 M HClO4 prior to the addition of 0.05 M H2O2, leading to the reduction of Pu(VI) and formation of a final 242Pu(V) stock (5 × 10−3 M). 238Pu and 242Pu concentrations were determined by liquid scintillation counting (LSC, Packard Tri-Carb TR2900) for all experiments.

Bacterial strain and growth conditions.

Pseudomonas sp. EPS-1W, previously isolated from the NNSS (36), was used in this study and is here referred to as wild-type EPS-1W. Cells were maintained on LB agar and grown in LB medium at 37°C with shaking (220 rpm) to late log phase before harvesting for experimental use. Late-log-phase cells were used to maximize cell biomass and minimize the potential release of cell debris into the EPS fraction due to cell lysis.

EPS removal and harvesting.

The method described in reference 37 was used to remove cell-bound EPS from the wild-type strain EPS-1W. Briefly, cells were cultured overnight in LB medium, pelleted and washed 3 times with Milli-Q H2O (>18 MΩ · cm), and suspended in a solution containing 2% Na3EDTA and 0.9% NaCl (pH 7) (37). After 24 h of incubation, the solution was centrifuged at 20,000 × g for a few minutes. Pelleted cells were washed 3 times with Milli-Q H2O prior to resuspension in 0.01 M NaCl and are here referred to as EPS-free cells.

Control experiments were performed to test the integrity of the EPS-free cells after the EPS removal process. After the last Milli-Q H2O wash, the optical density of the cell suspension was measured using a plate reader (Biotek) and cell integrity was examined by microscopy, including bright-field microscopy and DAPI (4′,6-diamidino-2-phenylindole; ThermoFisher Scientific) staining according to the manufacturer's instructions. Cells prior to EDTA incubation were included as the control. We found that the optical density did not change (see Fig. S1 in the supplemental material), and microscopic analysis revealed no detectable change in cell morphology before and after treatments (see Fig. S2 in the supplemental material), indicating that cell integrity was not compromised by the treatments. Based on these results, we concluded that our procedures did not cause significant cell lysis and therefore the extracted EPS are not likely to contain any significant contamination from cell debris. The Milli-Q H2O used has a pH of 6.0, well within the normal pH range in which Pseudomonas sp. EPS-1W grows.

Cell-bound EPS extracted from wild-type EPS-1W were isolated and purified using the ethanol precipitation method described in reference 38. Cold ethanol (final concentration, 50%) was mixed with the EPS solution and incubated for 24 h at 4°C before centrifugation at 5,000 × g for 30 min. The precipitated solid material was loaded into 10-kDa dialysis membranes and dialyzed against H2O for 48 h. The dialyzed solution was transferred to a new centrifuge tube and centrifuged at 10,000 × g for 2 h. The EPS precipitate was either resuspended in Milli-Q H2O or air dried for further characterization.

Pu interaction with cells.

In each set of sorption experiments, washed cells (wild type with intact EPS or EPS free) were incubated with Pu(IV) or Pu(V) in 0.01 M NaClO4 solution (pH 6.5) at room temperature with gentle rocking. The first set of cell sorption experiments were conducted with ∼10−10 M 238Pu(IV) or 238Pu(V) and ∼0.1 gcell/liter (all cell concentrations are based on cell dry weight, which was determined by washing cells in Milli-Q H2O, drying at 60°C overnight, and weighing using an analytical balance). At ∼1, 24, 48, and 72 h, a 3-ml cell suspension was centrifuged at 10,000 × g for 90 s, and the supernatant was sampled. Pu concentration in the supernatant was measured by LSC, and an aliquot of each sample was taken for LaF3 Pu(III, IV)/Pu(V, VI) oxidation state analysis. The second set of cell sorption experiments were conducted with ∼10−10 M 238Pu(IV) or 238Pu(V) and a range of cell concentrations (0.002 to 0.2 gcell/liter). After 24 h, a 3-ml cell suspension was centrifuged at 10,000 × g for 90 s, and the supernatant was sampled for Pu. A third set of cell sorption experiments were conducted specifically for transmission electron microscopy (TEM) analysis. In this case, much higher Pu concentrations were used, as described in detail below.

Pu interaction with purified EPS.

To quantify Pu(V) reduction rates in the presence of EPS, 4 × 10−9 238Pu(V) and 8 × 10−4 M 242Pu(V) were spiked into 1- to 100-ppm purified EPS solutions prepared from a 250-ppm EPS stock. Aliquots were taken at specific time intervals and centrifuged at 10,000 × g for 90 s, and the supernatant was analyzed for Pu concentration and Pu(III, IV)/Pu(V, VI) oxidation state.

Chemical analysis of EPS.

For a total protein content measurement, 12% trichloroacetic acid (TCA) was added to the EPS solution and the mixture was incubated on ice for 30 min before centrifugation at 15,000 × g for 20 min (36). The TCA precipitates were washed twice with 10 ml acetone and resuspended in 2 ml of 2-(N-morpholino)ethanesulfonic acid buffer (MES; 20 mM, pH 5.0), and the protein content was measured using the Bradford assay (Bio-Rad, Hercules, CA) with bovine serum albumin as a calibration standard. Total DNA content in the EPS solution was measured directly using pico-green (Life Technologies, California). Total carbohydrate content determination and glycosyl composition analysis were performed using gas chromatography-mass spectrometry (39) by the Complex Carbohydrate Research Center at the University of Georgia.

EPS reducing power was determined using the published method by Manivasagan et al. (40). Briefly, EPS (0.069 to 0.42 g/liter) was dissolved in phosphate buffer (pH 7) and equilibrated with 1% K3Fe(CN)3 for 20 min at 50°C. Ten percent TCA was added before samples were centrifuged at 5,000 × g for 30 min. The supernatant was collected and mixed with 0.25% FeCl3. After 60 min of incubation, absorbance at 700 nm was measured (Varian Cary 500 UV-Vis spectrometer). Milli-Q H2O was used as a negative control, and ascorbic acid was used as a positive control.

NMR analysis of EPS.

Dried EPS was loaded into clean nuclear magnetic resonance (NMR) tubes. The 13C{1H} cross polarization magic angle spinning (CP-MAS) spectrum was collected using a 600-MHz Bruker Avance III with a Bruker Very Fast MAS probe configured for 1.3-mm (outside diameter) rotors. Data collection conditions include 1 ms of contact time, a 2-s pulse delay, and a spinning rate of 60 kHz. The CP match condition was optimized on an external sample of uniformly labeled phenylalanine. After acquisition, the time domain data were first processed according to using a Cadzow enhancement procedure in order to enhance the signal-to-noise ratio of the processed NMR spectra. These methods have been shown to substantially improve the signal-to-noise ratio while retaining all the relevant spectral features (41).

TEM of cells.

TEM samples were prepared by equilibrating 242Pu(IV) or 242Pu(V) with a cell suspension (0.3 gcell/liter) in 0.01 M NaClO4 at pH 6.5. Pu concentrations used in TEM sample preparation are listed in Table 1. After 72 h of gentle agitation, cells were pelleted by centrifuging at 10,000 × g for 90 s and then washed three times with Milli-Q H2O and fixed for 1 h in 5% paraformaldehyde. The fixed cells were rinsed three times and suspended in Milli-Q H2O. The amount of Pu in the supernatant and the quantity removed during the washing steps were determined by LSC (Table 1). A 10-μl droplet of the cell suspension was placed on a Cu grid and air dried before TEM analysis on a Philips CM300 FEG super-twin microscope operated at 300 kV.

TABLE 1.

Sorption of Pu(IV) and Pu(V) to cells for TEM analysis

Target oxidation state Cell type Putotal (mol/liter) Pu(III, IV)initial (%) Pu sorbed (%) Pu removed during cell washes (%)
Pu(IV) EPS free 7.0 × 10−5 98 96 10
Wild type 5.0 × 10−5 98 92 9
Wild type 9.0 × 10−4 99 75 50
Pu(V) Wild type 7.0 × 10−5 10 98 3
Wild type 5.0 × 10−4 12 92 5
Open in a new tab

RESULTS

Cellular Pu sorption and reduction.

Pseudomonas sp. EPS-1W was used in this study as a model organism. We have focused on the cell-bound EPS because, as the outermost layer of the cell, the bound EPS are likely to play a primary role in Pu-cell interaction. Figure 1 shows the sorption kinetics of Pu(IV) and Pu(V), both present at ∼10−10 M, on wild type and EPS-free cells. Rapid initial sorption was observed for Pu(IV) in the presence and absence of cell-bound EPS. The sorption kinetics and affinity of Pu(IV) for the wild-type and EPS-free cells were similar, with over 60% of the Pu removed from solution after 3 days. In contrast to Pu(IV), an initially fast Pu(V) sorption to the wild-type cells (40%) was followed by a slow increase in sorption that reached equilibrium after 24 h (Fig. 1). Pu(V) sorption to the EPS-free cells was lower (20%), with no additional sorption observed over the course of the experiment (72 h). These data indicate that for Pu(V), wild-type cells are more effective at sorbing Pu than EPS-free cells.

FIG 1.

FIG 1

Open in a new tab

Sorption of Pu(IV) (top) and Pu(V) (bottom) to wild-type and EPS-free Pseudomonas sp. EPS-1W as a function of time (10−10 M Pu; pH 6.5). Error bars represent 2σ error calculated from triplicate samples.

To further evaluate Pu sorption to EPS-1W, the sorption of Pu (∼10−10 M) was measured as a function of cell concentration. Pu(IV) sorption increased with cell density, with maximum sorption (∼80%) occurring at 0.2 gcell/liter (see Fig. S3 in the supplemental material). This behavior suggests that Pu(IV) is removed from solution as a result of its association with cells. Additionally, the sorption of Pu(IV) to wild-type cells was similar to the sorption to EPS-free cells over the cell density range tested (0.002 to 0.2 gcell/liter). This indicates that the strong association of Pu(IV) with cells exists regardless of the presence or absence of cell-bound EPS.

In the case of Pu(V), sorption to wild-type cells followed a trend similar to that of Pu(IV), increasing with cell density to a maximum (∼80%) at 0.2 gcell/liter. However, the amount of Pu(V) sorbed to EPS-free cells was significantly reduced. While there is still a positive correlation between Pu(V) sorbed and cell density, the overall Pu(V) sorption was limited to ∼20% in the absence of cell-bound EPS. These results demonstrate that removal of cell-bound EPS significantly limits the cell surface binding of Pu(V). While both cell-bound EPS and cell wall exhibit chemical and/or physical attributes that promote binding of Pu(IV), cell-bound EPS are the primary contributor to Pu(V) sorption.

To understand the difference in Pu(V) interaction with wild-type and EPS-free cells, we determined the oxidation state of Pu remaining in the aqueous phase during the cell sorption experiments. With the wild-type cells, the oxidation state of the soluble Pu(V) rapidly reduced to Pu(III, IV), with complete reduction in solution observed by 48 h (Fig. 2). In contrast, there was no significant reduction of Pu(V) in solution with EPS-free cells over the 72 h of incubation period studied. The reduction of Pu(V) to Pu(III, IV) observed in the aqueous phase with the wild-type cells could have resulted from reduction by a small amount of EPS in wild-type cells that may have shed into the solution or from a Pu species reduced on the cell surface that was subsequently desorbed. Our data cannot distinguish between these or other possible mechanisms. Nevertheless, it is clear that the presence of EPS leads to the reduction of Pu(V).

FIG 2.

FIG 2

Open in a new tab

Fraction of Pu in the Pu(V, VI) oxidation state in solution during Pu(V) sorption to wild-type and EPS-free cells. The 0.08 gcell/liter data (shown here) are similar to those at other cell densities (see Fig. S3 in the supplemental material).

Reduction of Pu(V) by purified EPS.

The reduction of Pu(V) by EPS in the absence of cells was investigated using purified EPS from EPS-1W. As a function of EPS concentration, we assayed the Pu oxidation state in solution with the purified EPS (Fig. 3). Near-complete reduction of 4 × 10−9 M Pu(V) was observed at EPS concentrations as low as 1 ppm. Some reduction of Pu(V) to Pu(III, IV) (10 to 20%) in the EPS-free control samples was observed over the first time points but remained steady over the course of the experiments. The similar Pu(V) reduction levels by purified EPS and by the wild-type cells suggest that the underlying reduction and sorption mechanisms are likely the same. At a higher concentration of Pu(V) (8 × 10−4 M), near-complete reduction of Pu(V) was also observed with purified EPS at concentrations ranging from 10 to 25 ppm. Reducing power analysis based on ferricyanide reduction revealed the strong reducing power of the purified EPS, comparable to that of ascorbic acid (see Fig. S4 in the supplemental material).

FIG 3.

FIG 3

Open in a new tab

Reduction of 4 × 10−9 M 238Pu(V) (top) and 8 × 10−4 M 242Pu(V) (bottom) by EPS as a function of time (pH 6.5; 0.01 M NaClO4). Error bars represent 2σ error calculated from triplicate measurements.

As a first step toward identifying the EPS functional groups that control Pu(V) reduction, we performed compositional analysis of the purified EPS. Carbohydrates account for 75% of the total dry mass, with DNA and proteins accounting for 1% and 4%, respectively (the remaining 20% was not identified). NMR analysis was performed to identify functional groups associated with the EPS. The NMR solid-state 13C{1H} CP-MAS spectrum includes distinctive peaks corresponding to carboxylic acid and acetyl functional groups likely from peptides (see Fig. S5 in the supplemental material). Additional research is needed to identify which of the identified components of EPS can facilitate Pu(V) reduction. This is the subject of current efforts in our laboratory.

TEM analysis of Pu bound to Pseudomonas sp.

We examined the morphology and structure of Pu sorbed to the cell surface of wild-type and EPS-free cells using TEM. Sample preparation for the TEM analysis is summarized in Table 1. At the highest Pu concentrations investigated (∼10−4 M), the morphology and structure of Pu(IV) and Pu(V) sorbed to wild-type cells differed significantly (Fig. 4). Pu(IV) sorption experiments produced agglomerated nanocrystalline Pu(IV)O2 precipitates on the surface, consistent with the limited solubility of Pu(IV) under these conditions (Fig. 4A). High-resolution transmission electron microscopy (HRTEM) imaging (see Fig. S6 in the supplemental material) indicated that the nanocrystalline Pu(IV)O2 precipitates have a crystallite dimension of 2 to 3 nm, similar to those previously observed under similar solution conditions (13, 42, 43). The main features of the corresponding electron diffraction patterns match the face-centered cubic (fcc) Pu(IV)O2 crystal structure (see Fig. S6A in the supplemental material). However, the 2- to 3-nm dynamic range is near the limits of the TEM atomic resolution, resulting in broad diffraction rings compared to those typically observed for crystalline Pu(IV)O2.

FIG 4.

FIG 4

Open in a new tab

TEM images and EDS spectra of Pseudomonas sp. wild-type cells equilibrated with 9 × 10−4 M Pu(IV) (A) and 5 × 10−4 M Pu(V) (B), with the corresponding EDS spectra shown below (C and D, respectively). Pu was detectable on the cell surface in both cases, but nanocrystalline Pu precipitates formed only in the Pu(IV) sample (see Fig. S6 in the supplemental material).

In contrast to what was seen in the Pu(IV) experiment, nanocrystalline Pu(IV)O2 precipitates were not readily observed for the high-concentration Pu(V) sorption experiments (Fig. 4B). The Pu(V) sample concentration (7 × 10−4 M) was below the solubility limit of Pu(V)O2OH(am) (0.038 M) (44). However, upon reduction to Pu(IV), formation of nanocrystalline Pu(IV)O2 precipitates was expected. The energy-dispersive X-ray spectroscopy (EDS) spectrum unambiguously indicates the presence of Pu on the cell surface (Fig. 4D). HRTEM imaging suggests that some weakly ordered clusters may be present (see Fig. S6B in the supplemental material). However, in contrast to the Pu(IV) sorption experiments, Pu(V) reduced and sorbed by the bound EPS on the wild-type cells did not lead to the formation of nanocrystalline Pu(IV)O2 precipitates.

A comparison of Pu(IV) sorbed to wild-type versus EPS-free cells was performed using the lower-concentration (5 × 10−5 M) Pu(IV) TEM samples (Fig. 5). At this concentration, Pu(IV) is still orders of magnitude above its solubility. Nevertheless, no nanocrystalline Pu(IV)O2 precipitates were observed on either the wild-type or EPS-free cells. Similar to what was seen with the higher-concentration Pu(V) sample, HRTEM images suggested that some weakly ordered clusters may be present (see Fig. S7 in the supplemental material). Nevertheless, the formation of nanocrystalline Pu(IV)O2 precipitates appears to be inhibited by the presence of the cells and the EPS. The absence of nanocrystalline Pu(IV)O2 precipitates on wild-type and EPS-free cells is likely the result of monomeric Pu-EPS complexes and/or cell wall sorption.

FIG 5.

FIG 5

Open in a new tab

TEM images and EDS spectra of 5 × 10−5 M Pu(IV) equilibrated with wild-type cells (A) and 7 × 10−5 M Pu(IV) equilibrated with EPS-free cells (B). The corresponding EDS spectra are shown below (C and D, respectively). Pu was detected on the cell surface in both samples, but no nanocrystalline Pu precipitates were observed in either case (see Fig. S7 in the supplemental material).

DISCUSSION

We observed that Pu exhibited an affinity for Pseudomonas sp. strain EPS-1W, a common environmental bacterium previously isolated from the Pu-contaminated groundwater at the NNSS. Groundwater conditions at the NNSS are oxic, ∼pH 8, and of low ionic strength (45), similar to other contaminated sites, such as the Hanford Site in Washington and the Rocky Flats Environmental Technology Site in Colorado (46, 47). While our results are not directly linked to the Pu transport detected at the NNSS, our observations of Pu interaction with a common soil bacterium, Pseudomonas sp., suggest the potential importance of microbes and the specific role of EPS in facilitating redox transformation and sorption of Pu. The extent of the contribution of microbial activities to Pu transport at each site will likely depend on the identity and abundance of microorganisms (48), other competing geochemical processes (e.g., abiotic processes), and hydrologic factors.

Our study focused on the role of cell-bound EPS with regard to its direct contribution to Pu(V) reduction and Pu(IV) sorption. Pu(IV) sorbs quickly to both wild-type and EPS-free cells. However, Pu(V) sorption behavior is dependent on the presence of cell-bound EPS as depicted in Fig. 6. In the aqueous phase, oxidation state analyses indicate that EPS reduced Pu(V). The Pu(V) reduction kinetics in the presence of purified EPS indicate that even relatively small amounts of EPS can rapidly reduce Pu(V) to Pu(III, IV) at rates comparable to those observed in our wild-type cell sorption experiments. These results also indicate that the chemical reactivity of EPS does not appear to be altered by its removal from the cell surface.

FIG 6.

FIG 6

Open in a new tab

Model for Pu reduction and sorption by Pseudomonas sp. EPS-1W. For wild-type cells (left), Pu(V) reduction to Pu(IV), with subsequent cell surface sorption, is facilitated by the presence of bound EPS on the cell surface. This reduction has been confirmed by in vitro assay with purified EPS. For cells with bound EPS removed (right), reduction to Pu(IV) does not proceed and the amount of Pu(V) cell sorption is greatly decreased.

Detailed examination of Pu sorption behavior and precipitate morphology provides a conceptual model for the role of cell-bound EPS in Pu sorption, reduction, and precipitation processes. Based on the solubility of Pu(V)O2OH(am) (0.038 M) (44), precipitation of Pu(V) is not predicted to occur in any of our samples. Thus, only sorption and/or reduction is likely to control the behavior of Pu(V). Oxidation state analysis suggests that EPS facilitate Pu(V) reduction, which is consistent with an earlier examination of the role of bound EPS on U(VI) sorption by Shewanella under anoxic conditions (29). Thus, the sorption of Pu(V) to EPS can be described by a combination of reduction and sorption processes. Based on the solubility of Pu(IV)O2(am,hyd) (10−10.3 ± 0.5 M using the Nuclear Energy Agency Thermodynamic Database [44] or 10−10.4 ± 0.5 M using the more recent solubility estimates of Neck et al. [4]), precipitation of Pu(IV) may occur in our samples. However, sorption of 10−10 M Pu(IV) correlated with cell concentration (see Fig. S3 in the supplemental material), arguing against a pure precipitation mechanism at this concentration. Rather, our data favor Pu(IV) sorption to cell surface/EPS functional groups as a primary sorption mechanism for Pu(IV). Finally, TEM of Pu(IV) and Pu(V) sorbed to cells at high Pu concentrations yield distinct morphological features. Reduction of Pu(V) by bound EPS leads to sorption of monomeric Pu or amorphous Pu clusters on the cell surface. In contrast, Pu(IV) sorption leads to the formation of nanocrystalline Pu(IV)O2 precipitates at high concentrations and the formation of monomeric Pu or amorphous Pu clusters at lower concentrations. The strong influence of EPS on Pu(V) reduction, Pu(IV) sorption, and Pu(IV)O2 precipitate formation suggests that its presence (and the presence of its constituent biomolecules) in the environment may impact the biogeochemical cycling of Pu.

Follow-up work is needed to elucidate the specific mechanism of Pu(V) reduction by EPS. Carboxylic acid functional groups are strong metal complexants and could be involved in Pu complexation. Moreover, in some cases, they may lead to the reduction of Pu(V) (49). Acetyl groups are often found in EPS and on cell walls and have been attributed to UO22+ sorption by Pseudomonas sp. strain EPS-5028 (50). The presence of arabinose and mannose in the cell-bound EPS as revealed by the glycolysis analysis (see Table S1 in the supplemental material) may explain this high reducing power, given that these monosaccharides have been linked to the reductive capabilities of EPS purified from other bacteria (51). Interestingly, the fast reduction rate of Pu by EPS contrasts with the much slower kinetics and efficacy of Pu reduction by humic acids (52). Fast reduction kinetics by EPS have also been observed with UO22+, among other metals, and has been attributed to the presence of redox-active proteins (e.g., c-type cytochromes) present in EPS (32, 53, 54) of cells grown under anoxic conditions. These studies show that various components of EPS may contribute to Pu(V) reduction, highlighting the need for further analysis to determine which components are actively involved.

In conclusion, it appears that EPS may play an important role in the redox cycling of Pu and thus affect its mobility in the environment. Many open questions remain as to how Pu behavior is affected when microorganisms and their associated ligands coexist with inorganic minerals and other organic ligands in multicomponent groundwater systems. Nevertheless, in efforts to develop conceptual models to ascertain the dominant biogeochemical mechanisms controlling Pu mobility/immobility in the environment, it is evident that we cannot ignore the role of microorganisms.

Supplementary Material

Supplemental material
supp_82_24_7093__index.html (1.4KB, html)

ACKNOWLEDGMENTS

This work was prepared by LLNL under contract DEAC52-07NA27344.

We thank Parastoo Azadi at the Complex Carbohydrate Research Center at the University of Georgia for performing the glycosyl composition analysis.

Funding Statement

This work, including the efforts of Mark Boggs, Mavrik Zavarin, Yongqin Jiao, Zurong Dai, and Annie Kersting, was funded by the Subsurface Biogeochemical Research Program of the U.S. Department of Energy’s Office of Biological and Environmental Research, contract number SCW1053.

Footnotes

Supplemental material for this article may be found at http://dx.doi.org/10.1128/AEM.02572-16.

REFERENCES

  • 1.Ewing RC, Runde W, Albrecht-Schmitt TE. 2010. Environmental impact of the nuclear fuel cycle: fate of actinides. MRS Bull 35:859–866. doi: 10.1557/mrs2010.712. [DOI] [Google Scholar]
  • 2.Albright D, Kramer K. 2004. Fissile material—stockpiles still growing. Bull At Sci 60:14–16. [Google Scholar]
  • 3.Ewing RC. 1999. Nuclear waste forms for actinides. Proc Natl Acad Sci U S A 96:3432–3439. doi: 10.1073/pnas.96.7.3432. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Neck V, Altmaier M, Fanghanel T. 2007. Solubility of plutonium hydroxides/hydrous oxides under reducing conditions and in the presence of oxygen. C R Chim 10:959–977. doi: 10.1016/j.crci.2007.02.011. [DOI] [Google Scholar]
  • 5.Begg JD, Zavarin M, Zhao PH, Tumey SJ, Powell B, Kersting AB. 2013. Pu(V) and Pu(IV) sorption to montmorillonite. Environ Sci Technol 47:5146–5153. doi: 10.1021/es305257s. [DOI] [PubMed] [Google Scholar]
  • 6.Zavarin M, Roberts SK, Hakem N, Sawvel AM, Kersting AB. 2005. Eu(III), Sm(III), Np(V), Pu(V), and Pu(IV) sorption to calcite. Radiochim Acta 93:93–102. doi: 10.1524/ract.93.2.93.59413. [DOI] [Google Scholar]
  • 7.Banik NL, Buda RA, Burger S, Kratz JV, Trautmann N. 2007. Speciation and interactions of plutonium with humic substances and kaolinite in aquifer systems. J Alloys Compd 444:522–525. [Google Scholar]
  • 8.Choppin GR. 2003. Actinide speciation in the environment. Radiochim Acta 91:645–649. doi: 10.1524/ract.91.11.645.23469. [DOI] [Google Scholar]
  • 9.Kaplan DI, Powell BA, Duff MC, Demirkanli DI, Denham M, Fjeld RA, Molz FJ. 2007. Influence of sources on plutonium mobility and oxidation state transformations in vadose zone sediments. Environ Sci Technol 41:7417–7423. doi: 10.1021/es0706302. [DOI] [PubMed] [Google Scholar]
  • 10.Silva RJ, Nitsche H. 1995. Actinide environmental chemistry. Radiochim Acta 70-71:377–396. [Google Scholar]
  • 11.Kersting AB. 2013. Plutonium transport in the environment. Inorg Chem 52:3533–3546. doi: 10.1021/ic3018908. [DOI] [PubMed] [Google Scholar]
  • 12.Neck V, Altmaier M, Seibert A, Yun JI, Marquardt CM, Fanghanel T. 2007. Solubility and redox reactions of Pu(IV) hydrous oxide: evidence for the formation of PuO2+x(s, hyd). Radiochim Acta 95:193–207. [Google Scholar]
  • 13.Powell BA, Dai ZR, Zavarin M, Zhao PH, Kersting AB. 2011. Stabilization of plutonium nano-colloids by epitaxial distortion on mineral surfaces. Environ Sci Technol 45:2698–2703. doi: 10.1021/es1033487. [DOI] [PubMed] [Google Scholar]
  • 14.Fredrickson JK, Zachara JM, Balkwill DL, Kennedy D, Li SMW, Kostandarithes HM, Daly MJ, Romine MF, Brockman FJ. 2004. Geomicrobiology of high-level nuclear waste-contaminated vadose sediments at the Hanford Site, Washington State. Appl Environ Microbiol 70:4230–4241. doi: 10.1128/AEM.70.7.4230-4241.2004. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Francis AJ, Dobbs S, Nine BJ. 1980. Microbial activity of trench leachates from shallow-land, low-level, radioactive-waste disposal sites. Appl Environ Microbiol 40:108–113. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 16.Cherkouk A, Law GTW, Rizoulis A, Law K, Renshaw JC, Morris K, Livens FR, Lloyd JR. 2016. Influence of riboflavin on the reduction of radionuclides by Shewanella oneidensis MR-1. Dalton Trans 45:5030–5037. doi: 10.1039/C4DT02929A. [DOI] [PubMed] [Google Scholar]
  • 17.Lukšienė B, Druteikienė R, Pečiulytė D, Baltrūnas D, Remeikis V, Paškevičius A. 2012. Effect of microorganisms on the plutonium oxidation states. Appl Radiat Isot 70:442–449. doi: 10.1016/j.apradiso.2011.11.016. [DOI] [PubMed] [Google Scholar]
  • 18.Neu MP, Boukhalfa H, Merroun ML. 2010. Biomineralization and biotransformations of actinide materials. MRS Bull 35:849–857. doi: 10.1557/mrs2010.711. [DOI] [Google Scholar]
  • 19.Neu MP, Icopini GA, Boukhalfa H. 2005. Plutonium speciation affected by environmental bacteria. Radiochim Acta 93:705–714. [Google Scholar]
  • 20.Panak PJ, Nitsche H. 2001. Interaction of aerobic soil bacteria with plutonium(VI). Radiochim Acta 89:499–504. [Google Scholar]
  • 21.Wildung RE, Garland TR. 1982. Effects of plutonium on soil microorganisms. Appl Environ Microbiol 43:418–423. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22.Panak PJ, Booth CH, Caulder DL, Bucher JJ, Shuh DK, Nitsche H. 2002. X-ray absorption fine structure spectroscopy of plutonium complexes with Bacillus sphaericus. Radiochim Acta 90:315–321. [Google Scholar]
  • 23.Ohnuki T, Yoshida T, Ozaki T, Kozai N, Sakamoto F, Nankawa T, Suzuki Y, Francis AJ. 2009. Modeling of the interaction of Pu(VI) with the mixture of microorganism and clay. J Nucl Sci Technol 46:55–59. doi: 10.1080/18811248.2007.9711507. [DOI] [Google Scholar]
  • 24.Rusin PA, Quintana L, Brainard JR, Strietelmeier BA, Tait CD, Ekberg SA, Palmer PD, Newton TW, Clark DL. 1994. Solubilization of plutonium hydrous oxide by iron-reducing bacteria. Environ Sci Technol 28:1686–1690. doi: 10.1021/es00058a021. [DOI] [PubMed] [Google Scholar]
  • 25.Icopini GA, Lack JG, Hersman LE, Neu MP, Boukhalfa H. 2009. Plutonium(V/VI) reduction by the metal-reducing bacteria Geobacter metallireducens GS-15 and Shewanella oneidensis MR-1. Appl Environ Microbiol 75:3641–3647. doi: 10.1128/AEM.00022-09. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Boukhalfa H, Icopini GA, Reilly SD, Neu MP. 2007. Plutonium(IV) reduction by the metal-reducing bacteria Geobacter metallireducens GS15 and Shewanella oneidensis MR1. Appl Environ Microbiol 73:5897–5903. doi: 10.1128/AEM.00747-07. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Plymale AE, Bailey VL, Fredrickson JK, Heald SM, Buck EC, Shi L, Wang Z, Resch CT, Moore DA, Bolton H. 2012. Biotic and abiotic reduction and solubilization of Pu(IV)O2·xH2O(am) as affected by anthraquinone-2,6-disulfonate (AQDS) and ethylenediaminetetraacetate (EDTA). Environ Sci Technol 46:2132–2140. doi: 10.1021/es2030752. [DOI] [PubMed] [Google Scholar]
  • 28.Renshaw JC, Law N, Geissler A, Livens FR, Lloyd JR. 2009. Impact of the Fe(III)-reducing bacteria Geobacter sulfurreducens and Shewanella oneidensis on the speciation of plutonium. Biogeochemistry 94:191–196. doi: 10.1007/s10533-009-9318-8. [DOI] [Google Scholar]
  • 29.Cao B, Ahmed B, Kennedy DW, Wang Z, Shi L, Marshall MJ, Fredrickson JK, Isern NG, Majors PD, Beyenal H. 2011. Contribution of extracellular polymeric substances from Shewanella sp. HRCR-1 biofilms to U(VI) immobilization. Environ Sci Technol 45:5483–5490. doi: 10.1021/es200095j. [DOI] [PubMed] [Google Scholar]
  • 30.Deo RP, Songkasiri W, Rittmann BE, Reed DT. 2010. Surface complexation of neptunium(V) onto whole cells and cell components of Shewanella alga: modeling and experimental study. Environ Sci Technol 44:4930–4935. doi: 10.1021/es9035336. [DOI] [PubMed] [Google Scholar]
  • 31.Marshall MJ, Beliaev AS, Dohnalkova AC, Kennedy DW, Shi L, Wang Z, Boyanov MI, Lai B, Kemner KM, McLean JS, Reed SB, Culley DE, Bailey VL, Simonson CJ, Saffarini DA, Romine MF, Zachara JM, Fredrickson JK. 2006. c-Type cytochrome-dependent formation of U(IV) nanoparticles by Shewanella oneidensis. PLoS Biol 4:e268. doi: 10.1371/journal.pbio.0040268. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32.Stylo M, Alessi DS, Shao PP, Lezama-Pacheco JS, Bargar JR, Bernier-Latmani R. 2013. Biogeochemical controls on the product of microbial U(VI) reduction. Environ Sci Technol 47:12351–12358. doi: 10.1021/es402631w. [DOI] [PubMed] [Google Scholar]
  • 33.Kobashi A, Choppin GR, Morse JW. 1988. A study of techniques for separating plutonium in different oxidation states. Radiochim Acta 43:211–215. [Google Scholar]
  • 34.Orlandini KA, Penrose WR, Nelson DM. 1986. Pu(V) as the stable form of oxidized plutonium in natural-waters. Mar Chem 18:49–57. doi: 10.1016/0304-4203(86)90075-7. [DOI] [Google Scholar]
  • 35.Cohen D. 1961. Electrochemical studies of plutonium ions in perchloric acid solution. J Inorg Nucl Chem 18:207–210. doi: 10.1016/0022-1902(61)80389-8. [DOI] [Google Scholar]
  • 36.Moser D, Hamilton-Brehm S, Fisher J, Bruckner J, Kruger B, Sackett J, Russel CE, Onstott TC, Czerwinski K, Zavarin M, Campbell JH. 2014. Final report: radiochemically-supported microbial communities: a potential mechanism for biocolloid production of importance to actinide transport. Desert Research Institute, Las Vegas, NV. [Google Scholar]
  • 37.Xu C, Santschi PH, Schwehr KA, Hung CC. 2009. Optimized isolation procedure for obtaining strongly actinide binding exopolymeric substances (EPS) from two bacteria (Sagittula stellata and Pseudomonas fluorescens Biovar II). Bioresour Technol 100:6010–6021. doi: 10.1016/j.biortech.2009.06.008. [DOI] [PubMed] [Google Scholar]
  • 38.Cao B, Shi L, Brown RN, Xiong Y, Fredrickson JK, Romine MF, Marshall MJ, Lipton MS, Beyenal H. 2011. Extracellular polymeric substances from Shewanella sp. HRCR-1 biofilms: characterization by infrared spectroscopy and proteomics. Environ Microbiol 13:1018–1031. doi: 10.1111/j.1462-2920.2010.02407.x. [DOI] [PubMed] [Google Scholar]
  • 39.Jiao Y, Cody GD, Harding AK, Wilmes P, Schrenk M, Wheeler KE, Banfield JF, Thelen MP. 2010. Characterization of extracellular polymeric substances from acidophilic microbial biofilms. Appl Environ Microbiol 76:2916–2922. doi: 10.1128/AEM.02289-09. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40.Manivasagan P, Sivasankar P, Venkatesan J, Senthilkumar K, Sivakumar K, Kim SK. 2013. Production and characterization of an extracellular polysaccharide from Streptomyces violaceus MM72. Int J Biol Macromol 59:29–38. doi: 10.1016/j.ijbiomac.2013.04.012. [DOI] [PubMed] [Google Scholar]
  • 41.Man P, Bonhomme C, Babonneau F. 2014. Denoising NMR time-domain signal by singular-value decomposition accelerated by graphics processing units. Solid State Nucl Magn Reson 61-62:28–34. [DOI] [PubMed] [Google Scholar]
  • 42.Zavarin M, Zhao P, Dai Z, Kersting AB. 2014. Plutonium sorption and precipitation in the presence of goethite at 25 and 80 degrees C. Radiochim Acta 102:983–997. [Google Scholar]
  • 43.Romanchuk AY, Kalmykov SN, Egorov AV, Zubavichus YV, Shiryaev AA, Batuk ON, Conradson SD, Pankratov DA, Presnyakov IA. 2013. Formation of crystalline PuO2+x·nH2O nanoparticles upon sorption of Pu(V,VI) onto hematite. Geochim Cosmochim Acta 121:29–40. doi: 10.1016/j.gca.2013.07.016. [DOI] [Google Scholar]
  • 44.Guillaumont R, Fanghanel T, Fuger J, Neck V, Palmer D. 2003. Update on the chemical thermodynamics of uranium, neptunium, plutonium, americium and technetium, vol 5 OECD Nuclear Energy Agency, Issy-les-Moulineaux, France. [Google Scholar]
  • 45.Kersting AB, Efurd DW, Finnegan DL, Rokop DJ, Smith DK, Thompson JL. 1999. Migration of plutonium in ground water at the Nevada Test Site. Nature 397:56–59. doi: 10.1038/16231. [DOI] [Google Scholar]
  • 46.Dai MH, Buesseler K, Pike SM. 2005. Plutonium in groundwater at the 100K-Area of the US DOE Hanford site. J Contam Hydrol 76:167–189. doi: 10.1016/j.jconhyd.2004.08.004. [DOI] [PubMed] [Google Scholar]
  • 47.Santschi PH, Roberts KA, Guo LD. 2002. Organic nature of colloidal actinides transported in surface water environments. Environ Sci Technol 36:3711–3719. doi: 10.1021/es0112588. [DOI] [PubMed] [Google Scholar]
  • 48.Kieft TL, Amy PS, Brockman FJ, Fredrickson JK, Bjornstad BN, Rosacker LL. 1993. Microbial abundance and activities in relation to water potential in the vadose zone of arid and semiarid sites. Microb Ecol 26:59–78. doi: 10.1007/BF00166030. [DOI] [PubMed] [Google Scholar]
  • 49.Reed DT, Wygmans DG, Aase SB, Banaszak JE. 1997. Reduction of Np(VI) and Pu(VI) by organics chelating agents. Radiochim Acta 82:109–114. [Google Scholar]
  • 50.Marques AM, Bonet R, Simonpujol MD, Fuste MC, Congregado F. 1990. Removal of uranium by an exopolysaccharide from Pseudomonas sp. Appl Microbiol Biotechnol 34:429–431. [Google Scholar]
  • 51.Kang FX, Alvarez PJ, Zhu DQ. 2014. Microbial extracellular polymeric substances reduce Ag+ to silver nanoparticles and antagonize bactericidal activity. Environ Sci Technol 48:316–322. doi: 10.1021/es403796x. [DOI] [PubMed] [Google Scholar]
  • 52.Andre C, Choppin GR. 2000. Reduction of Pu(V) by humic acid. Radiochim Acta 88:613–616. [Google Scholar]
  • 53.Merroun ML, Selenska-Pobell S. 2008. Bacterial interactions with uranium: an environmental perspective. J Contam Hydrol 102:285–295. doi: 10.1016/j.jconhyd.2008.09.019. [DOI] [PubMed] [Google Scholar]
  • 54.Gorby YA, Lovley DR. 1992. Enzymatic uranium precipitation. Environ Sci Technol 26:205–207. doi: 10.1021/es00025a026. [DOI] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplemental material
supp_82_24_7093__index.html (1.4KB, html)
AEM.02572-16_zam999117552so1.pdf (815.7KB, pdf)

Articles from Applied and Environmental Microbiology are provided here courtesy of American Society for Microbiology (ASM)

RESOURCES